A Key Role for Cc Chemokine Receptor 4 in Lipopolysaccharide-Induced Endotoxic Shock

Chemokines are a superfamily of small proteins involved both in routine leukocyte trafficking and in the activation and recruitment of specific cell populations to sites of inflammation and infection. There are four classes of chemokines, named according to the spacing of the first two of a conserved four-cysteine motif: CC, CXC, C, and CX₃C. The majority of chemokines belong to the CC and CXC subfamilies. For the C and CX₃C subfamilies, only one member has been identified to date, lymphotactin and fractalkine (or neurotactin), respectively 1,2. The biological activity of chemokines is mediated by their interaction with a family of seven-transmembrane G protein–coupled receptors. To date, there are published reports for five CXC chemokine–specific receptors, nine CC, a CX₃C, and a C chemokine receptor as well as the Duffy antigen and several virally encoded chemokine receptors 3,4. In vitro studies have shown that most chemokines are able to bind and activate more than one chemokine receptor, and the majority of chemokine receptors bind multiple ligands although there are exceptions. Yet in spite of this apparent redundancy, there are now increasing numbers of reports in the literature associating specific chemokine receptor–ligand pairs with particular disease states or tissue-specific lymphocyte homing. Much of this information has been obtained from studies of chemokine or chemokine receptor knockout mice.

We have previously reported the molecular cloning of both human and murine CC chemokine receptor (CCR) 4 5,6. Although originally identified as a receptor for macrophage inflammatory protein (MIP)-1α and regulated upon activation, normal T cell expressed and secreted chemokine (RANTES), CCR4 is in fact a high affinity receptor for two recently described CC chemokines, thymus and activation-regulated chemokine (TARC) 7 and macrophage-derived chemokine (MDC) 8. CCR4 mRNA is expressed predominantly in the thymus, spleen, and in peripheral blood leukocytes, including T cells, basophils, monocytes 5, macrophages, and platelets 9. In addition, several reports have shown that CCR4 is highly expressed by human T cells polarized towards the Th2 phenotype 10,11,12. These Th2 cells are also responsive to the CCR4 ligands TARC and MDC, leading investigators to suggest a role for CCR4 in the development of Th2 responses. Finally, a recent report has shown that CCR4 is also expressed on all skin homing T cells expressing cutaneous lymphocyte antigen (CLA) 13.

To study the role of CCR4 in vivo, we have generated CCR4-deficient mice by gene targeting. Here, we show that in spite of the reported high level expression of CCR4 on human Th2-polarized cells, CCR4 deletion had no effect on the development of the response in a classical Th2-dependent murine model of airway inflammation. However, CCR4^−/− mice showed an unexpected resistance to the lethal effects of LPS in two models of LPS-induced endotoxic shock.

The murine (m)CCR4 gene was isolated from an HM-1 embryonic stem (ES) cell library in λFIXII vector (Stratagene) by plaque hybridization using murine CCR4 cDNA as a probe 6. Two unique clones of 12.18 and 13.08 kb were shown to contain the mCCR4 coding sequence by PCR, using specific primers 6. An 8.5-kb fragment of genomic DNA confirmed to contain the CCR4 coding sequence by Southern blotting was subcloned into pBluescript II SK⁻ to generate pCCR4. The entire CCR4 coding sequence was then removed as an NheI-HpaI fragment and replaced with a neo cassette (derived from pMC1neoPolyA; CLONTECH Laboratories, Inc.). The resulting construct was digested with EcoRV and Eco47-3, and religated to generate a plasmid containing a long arm of homology of 4,904 bp and a short arm of homology of 1,318 bp. Finally, a thymidine kinase (tk) cassette was inserted into the HindIII-XhoI site of the plasmid to produce the targeting vector. The targeting vector was linearized with NotI and electroporated into HM-1 ES cells as described previously 14. Gancyclovir and G418-resistant clones were selected. DNA was isolated from resistant clones using DNAzol (GIBCO BRL), and the presence of the transgene was detected by PCR and verified by Southern hybridization on PstI-digested genomic DNA, using a 466-bp probe derived by AvrII-NsiI digestion of pCCR4. Seven independent transgene-containing ES cell clones were used to produce chimeric mice by blastocyst injection according to standard procedures 15. Two 100% chimeric males derived from two independent ES clones were mated with a 100% chimeric female to generate heterozygous CCR4^+/− mice, and littermates from the matings of CCR4^+/− mice were analyzed for the presence of homozygous CCR4^−/− mice by Southern blot analysis or by PCR on tail DNA. The sequences of the PCR primers used are: (primer 1, neomycin gene) 5′-CGCTTCCTCGTGCTTTACGGTAT; (primer 2, CCR4 3′ untranslated region) 5′-ATAGCCTTGGCTGGTCTGGAACTA; (primer 3, mCCR4 coding region sense primer) 5′-CCAAAGATGAATGCCACAGAGGTCACAG; (primer 4, mCCR4 coding region antisense primer) 5′-TTACAAAGCGTCACGGAAGTCATG.

Total RNA was isolated from thymocytes, splenocytes, or peritoneal lavage cells using Trizol™ (GIBCO BRL). 1 μg of total RNA from thymocytes and splenocytes or total RNA from 10⁶ peritoneal exudate cells was reverse transcribed using Superscript™ (GIBCO BRL) and oligo dT_12–18 primer according to the manufacturer's instructions. One twentieth of the cDNA synthesis reaction was then subjected to 25 cycles of PCR using AmpliTaq™ (PerkinElmer) and PCR primers based on the EMBL/GenBank/DDBJ database entries for MIP-2 (X53798), mMDC (AF052505), and mCCR4 (X90862). PCR products were analyzed on 1% agarose gels stained with ethidium bromide, and bands migrating at the correct molecular weight were verified by direct sequencing. For semiquantitative reverse transcriptase (RT)-PCR, bands were quantitated using Kodak Digital Science v1.0 software and results are expressed as arbitrary units of mRNA.

CD8⁺ and Ig⁺ depleted cells from lymph nodes and spleens of naive CCR4^+/+ and CCR4^−/− mice were cultured for 4 d on plates coated with an anti-CD3 antibody (145-2C11; BD PharMingen) as described previously 16 in the presence of either murine IL-12 (500 pg/ml; R&D Systems) plus anti–IL-4 mAb (10 μg/ml) or murine IL-4 (500 U/ml; ImmunoKontact) plus anti–IFN-γ mAb (10 μg/ml; BD PharMingen). After a 5-d culture period, cells were washed and restimulated for 24 h at a density of 2 × 10⁵ cells per well on an anti-CD3–coated 96-well plate (Costar) in the presence of murine IL-2 (50 U/ml; R&D Systems).

Homozygous CCR4^−/− mice were backcrossed with C57BL/6 mice (Centre d'Elevage Janvier) for four generations. Age- and weight-matched CCR4^−/− and CCR4^+/+ littermates from heterozygote (CCR4^+/−) matings from the fourth backcross were used in this study to control for strain background. Mice (20–25 g) of either sex were immunized intraperitoneally with 10 μg of OVA (A-5503; Sigma-Aldrich) in 0.2 ml of alum (Serva). Control mice received an injection of saline (0.9% wt/vol NaCl) alone. 14 d later, mice were anesthetized by inhaled 2% FORENE™ (Abbott) and 50 μg of OVA was administered to the lungs (in 50 μl of saline) intranasally as described previously 17. Control mice received 50 μl saline only. This procedure was repeated daily for 5 d. Animals were finally killed by lethal injection of 60 mg/kg pentobarbital. Student's t test was used for statistical analysis, except that for the analysis of the survival curves, we performed the log rank test (two-tailed).

Phenol-extracted bacterial LPS from Escherichia coli 055:B5 (List Biological Laboratories) was administered at 60, 90, and 120 mg/kg intraperitoneally for the high dose LPS shock model. For the low dose LPS shock model, mice received 1, 2, and 4 μg of LPS with 8 mg d-galactosamine (D-gal; Fluka) in 0.5 ml saline. Animals were killed by CO₂ asphyxiation at the time points indicated in the figures.

Bronchial hyperreactivity (BHR) was measured by recording respiratory pressure curves by whole body plethysmography 18 in response to inhaled methacholine (MCh; Sigma-Aldrich) using a Buxco^® apparatus (EMKA Technologies). The airway reactivity was expressed in enhanced pause (Penh) as described previously 19.

Blood (20 μl) was collected from the retroorbital plexus of mice into a heparinized micropipette, then transferred to a Unopette microcollection system for platelet determination (Becton Dickinson). Three independent samples of platelets were counted in a hemocytometer. A minimum of 100 cells were counted, and the arithmetic mean of the three counts was calculated 20.

OVA-specific IgM, IgG1, IgG2a, and IgE titers were measured in serum samples obtained 3 d after the final intranasal treatment with OVA using a standard ELISA protocol 19. Quantification of murine TNF-α in serum and in culture supernatant and IL-1β, IL-6, and MIP-1α in serum was determined using cytokine-specific ELISAs as per the manufacturer's protocol (R&D Systems).

Bronchoalveolar lavage (BAL) and peritoneal lavage cells were harvested and differential cell counts were performed on cytospin preparations stained with Diff-Quik™ (Baxter Diagnostics). A minimum of 200 cells were counted per field, with 3 fields per sample for BAL, and 5 fields per sample for peritoneal lavage.

For phenotypic analysis of peritoneal cells by flow cytometry, lavaged cells were resuspended at 10⁶ cells/ml in PBS containing 1% BSA and 0.01% azide (FACS buffer). Cells were incubated with Fc block (BD PharMingen) for 10 min at 4°C, washed twice with FACS buffer, then incubated for 20 min with FITC-labeled rat anti–mouse F4/80 antibody (Serotec). Cells were washed twice, resuspended in 200 μl FACS buffer, and analyzed on a Becton Dickinson FACScan™ flow cytometer with CELLQuest™ software.

Spleens from CCR4^+/+ and CCR4^−/− mice were dispersed through a 70-μm nylon cell strainer (Becton Dickinson). Erythrocytes were removed by hypotonic lysis. Cells were harvested by centrifugation and resuspended in RPMI medium containing 5% FCS and 2 mM glutamine. Chemotaxis assays were performed using the micro-Boyden chamber method with 5-μm filters 21. Recombinant human and mouse chemokines were purchased from R&D Systems or made in house.

The murine CCR4 gene was deleted through homologous recombination using the targeting vector shown in Fig. 1 a. Targeted ES cells were used to generate chimeric mice, which transmitted the transgene through the germline. Southern blot analysis confirmed that the CCR4 gene had been deleted using a DNA probe located in the 5′ noncoding region of the gene (Fig. 1 b). PCR was also used to confirm that the CCR4 coding sequence had indeed been deleted while confirming the presence of the neomycin transgene and 3′ untranslated sequence of CCR4. The CCR4 knockout mice were viable, appeared to develop normally, and showed no overt morphological or behavioral defects in the unstressed state. RT-PCR was used to demonstrate that the mRNA for CCR4 was not present in the thymus, spleen (Fig. 1 c), or peritoneal cells (Fig. 1 d) of targeted animals.

The ability of splenocytes and thymocytes, isolated from the targeted and wild-type mice, to migrate in response to the proposed CCR4 ligands was then assessed. Splenocytes (Fig. 1 e) and thymocytes (data not shown) from CCR4^−/− mice had no chemotactic response to TARC or MDC, whereas splenocytes from CCR4^+/+ mice responded with the characteristic dose–response curves, confirming that the gene deleted in this study is an endogenous TARC and MDC receptor. Surprisingly, splenocytes isolated from the CCR4^−/− mice did not respond to human MIP-1α (Fig. 1 e) or to murine MIP-1α (data not shown) in at least four independent experiments. However, the response to human RANTES was similar in both groups of mice (Fig. 1 e). Murine RANTES receptors described to date (CCR1, CCR5, and CCR3) have also been shown to bind and signal in response to MIP-1α in vitro. Yet RT-PCR analysis of the cells used in the study confirmed that deletion of the CCR4 gene did not interfere with expression of these other receptors (data not shown). This result suggested that in these cell types, CCR4 is a physiological receptor for MIP-1α. At present, we are unable to explain this astonishing result. However, most of the data on chemokine receptor ligand specificity has been obtained from in vitro studies. As such it is possible that in vivo, other factors affect chemokine receptor selectivity. Alternatively, our observations could reflect the need for cooperativity between distinct chemokine receptors to generate ligand specificity.

The role of CCR4 in the in vitro differentiation of T cells was addressed next. Naive CD4⁺ T cells were cultured for 4 d on plates coated with an anti-CD3 antibody either in the presence of IL-12 plus anti–IL-4 antibody, which induces Th1 cell differentiation, or in the presence of IL-4 plus anti–IFN-γ antibody, which induces Th2 cell differentiation. Cells were then restimulated with the anti-CD3 antibody for 24 h. CD4⁺ T cells initially cultured in the presence of IL-4 showed no significant difference in the production of IL-4 (CCR4^+/+ T cells, 105 ± 15 pg/ml; CCR4^−/− T cells, 190 ± 10 pg/ml) after restimulation. Likewise, T cells cultured with IL-12 produced comparable levels of IFN-γ (CCR4^+/+, 76 ± 19 ng/ml; CCR4^−/−, 49 ± 8 ng/ml). These results indicate that in vitro, Th2 and Th1 cell differentiation was not impaired in the CCR4^−/− mice. As in vitro–derived Th2 T cells have previously been shown to express CCR4, we tested whether the cells used in this study could respond to CCR4 ligands in chemotaxis assays. Cells derived from the CCR4^−/− mice failed to migrate in response to MDC, whereas cells derived from the CCR4^+/+ mice had a robust chemotactic response to MDC (Fig. 2 a). A similar result was obtained using TARC (data not shown). In addition, in vitro–derived Th2 T cells from both wild-type and CCR4^−/− mice responded to RANTES (Fig. 2 a).

The effect of the CCR4 deletion in vivo was studied using an OVA-induced murine model of airway inflammation, a predominantly Th2-associated response 22. Repeated intranasal OVA challenges of immunized CCR4^+/+ and CCR4^−/− littermates resulted in a significant increase in BHR in response to inhaled MCh (3 × 10⁻² M) compared with saline-challenged mice (Fig. 2 a). Penh values were 0.8 ± 0.1 in saline compared with 1.77 ± 0.2 in OVA-challenged CCR4^+/+ mice, and 0.64 ± 0.11 in saline compared with 1.88 ± 0.33 in OVA-challenged CCR4^−/− mice. Comparable OVA-induced eosinophilia was observed both in BALF (Fig. 2 b) and in lung tissue (data not shown) of CCR4^+/+ and CCR4^−/− littermates, a finding consistent with the induction of a Th2 response in the airways. In addition, no significant differences were observed in the BAL fluid (BALF) in either the total cell count or individual leukocyte populations (eosinophils, macrophages, lymphocytes, and neutrophils) between OVA-challenged CCR4^+/+ and CCR4^−/− littermates (Fig. 2 b). To confirm that efficient antigen priming had occurred in the periphery, serum titers of OVA-specific IgM, IgG1, IgG2a, and IgE were measured. Again, all OVA-specific Ig titers were comparable in CCR4^+/+ and CCR4^−/− littermates after OVA sensitization and challenge (Fig. 2 c). Taken together, our results suggest that deletion of the CCR4 gene does not impair the development of a Th2 response in vivo in this model. This is in contrast to the data recently reported by Gonzalo et al. 23, in which the administration of polyclonal antibodies against MDC, one of the ligands for CCR4, protected against eosinophilia and BHR. However, there are several explanations for the differences between the results. First, there may be an alternative receptor for MDC on activated cells. Second, the genetic background of mice used in each study may differentially influence susceptibility. Third, the polyclonal antibodies used may have depleted MDC binding cells. These hypotheses will be tested when specific mAbs against murine MDC become available.

CCR4 is also expressed on other cell types, such as platelets 9, monocytes 5,24, and macrophages (Parums, D., and Power, C.A., unpublished data; see also Fig. 1 d). Therefore, we assessed the effect of the CCR4 deletion during LPS-induced endotoxic shock, an inflammatory model in which these cells types have been implicated 25,26. LPS (60–120 mg/kg) was injected intraperitoneally into CCR4^+/+ and CCR4^−/− littermates and survival was assessed daily for 6 d (Fig. 3 a). All CCR4^+/+ mice died between 2 and 4 d after LPS injection. In contrast, 14 out of 15 CCR4^−/− mice were alive on day 6, demonstrating significant resistance to 60 mg/kg LPS (P < 0.001). Indeed, the CCR4^−/− mice were also strikingly resistant to LPS doses of up to 120 mg/kg. Control mice (saline-injected CCR4^−/− mice, n = 4, and CCR4^+/+ mice, n = 4) remained alive and healthy throughout the 6-d study (data not shown). Interestingly, during the first few hours after LPS administration, CCR4^−/− mice still showed signs of endotoxemia such as shivering and lethargy. However, these effects were visually milder than in the CCR4^+/+ mice.

Intraperitoneal injection of a high dose of LPS is followed by a marked thrombocytopenia and accumulation of platelets in the liver and spleen 27. Blood samples from CCR4^−/− and CCR4^+/+ mice contained similar numbers of platelets. Furthermore, in an independent experiment, a superimposable decrease in blood platelet count occurred in both CCR4^+/+ (n = 3) and CCR4^−/− mice (n = 3) in the first 20 h after injection of high dose LPS (Fig. 3 b), indicating that there was no obvious difference in platelet mobilization between the two groups. However the platelet count returned to normal in the CCR4^−/− mice by 5 d after treatment.

As a comparison, the effect of the CCR4 deletion in a low dose LPS endotoxic shock model was studied. In this model, the susceptibility of mice to a low doses of LPS (1, 2, and 4 μg) is enhanced by coinjection of 8 mg of D-gal 28. Within 24 h of intraperitoneal injection of the LPS and D-gal combination, only 1 in 12 of the CCR4^+/+ mice survived, whereas 9 out of 12 of the CCR4^−/− littermates survived when treated with 1 μg of LPS plus D-gal (P < 0.02; Fig. 3 c), also demonstrating increased resistance to low dose LPS. However, when administered at 4 μg of LPS with D-gal, all CCR4^−/− mice tested died but with a 6-h delay compared with the wild-type mice.

LPS is known to stimulate the release of proinflammatory cytokines such as TNF-α and IL-1β from monocytes, macrophages, and neutrophils 29. Therefore, the effect of the high dose LPS response was studied in more detail. A sharp increase in serum TNF-α levels was observed in CCR4^+/+ mice 1.5 h after LPS injection, which returned to baseline by 4 h after treatment. Interestingly, CCR4^−/− mice failed to induce significant levels of serum TNF-α after LPS injection (P < 0.002 at 1.5 h; Fig. 4 a). Cells isolated by peritoneal lavage from naive CCR4^−/− mice produced significantly lower levels of TNF-α when cultured for 18 h in the presence of LPS, compared with cells isolated from naive CCR4^+/+ mice (107 ± 11 compared with 201 ± 18 pg/ml, P < 0.0079; Fig. 4 b). In addition, a sixfold decrease in serum IL-1β in CCR4^−/− mice was observed 3 h after LPS injection (P < 0.002) compared with CCR4^+/+ mice (Fig. 4 c). This suggests that the observed resistance to LPS may in part be due to decreased TNF-α and IL-1β production, implying that CCR4 is indirectly involved in the production of these cytokines. In contrast, IL-6 production was unaltered (data not shown), suggesting that in CCR4^−/− mice the regulation of IL-6 can be independent from that of TNF-α and IL-1β. Further analysis of the CCR4^−/− samples also revealed a parallel decrease in serum MIP-1α levels (P < 0.0249 at 1.5 h; Fig. 4 d), pointing to a possible macrophage defect.

The cellular composition of the peritoneal lavage was therefore assessed at various times after injection of high dose LPS. No significant differences were seen in the total number of cells at early time points. At 24 h after high dose LPS treatment, the number of neutrophils detected in the lavage of both groups of mice was comparable (Fig. 5 a). This is as expected, since to date there is no evidence for the expression of CCR4 on neutrophils. Furthermore, we have shown that neutrophils isolated from the peritoneal cavity of wild-type mice after thioglycollate treatment do not respond to CCR4 ligands in chemotaxis assays. In addition, thioglycollate-elicited neutrophils from CCR4^−/− mice respond normally to MIP-1α and MIP-2 (our unpublished data). However, CCR4^−/− mice had significantly fewer macrophages than CCR4^+/+ mice (P < 0.0099; Fig. 5 b). These findings were consistent with the observed decrease in the mRNA expression of the macrophage-associated chemokines MIP-2 30 and MDC 31 (Fig. 5c and Fig. d). We confirmed the apparent differences in macrophage numbers seen on the cytospins by FACS^® analysis for macrophage markers. We found decreased numbers of CD11b⁺ and CD14⁺ cells (data not shown) and an absence or severely reduced number of F4/80-expressing cells in the CCR4^−/− mice compared with the CCR4^+/+ mice 24 h after LPS treatment (Fig. 5 f). However, F4/80⁺ cells were detected in naive mice (Fig. 5 e) as well as at earlier time points after LPS injection in the CCR4^−/− mice (data not shown). These results may imply a defect in the ability of CCR4^−/− mice to retain a particular macrophage population expressing F4/80, in the peritoneum, which in turn may confer LPS resistance in CCR4^−/− mice.

The F4/80 antigen is an unusual seven-transmembrane receptor in that its extracellular domain is composed of EGF domain repeats. Its ligand(s) and precise function remain unknown 32, but while it is well established that activated macrophages express F4/80, a population of CD5⁺ peritoneal B cells has also recently been shown to express this marker 33. Interestingly, we also noted a decrease in the number of lymphocytes in the peritoneal lavage 24 h after high dose LPS treatment in CCR4^−/− mice (results not shown). However, the identity of the affected lymphocyte population and its contribution, if any, to the mechanism of LPS resistance in CCR4^−/− mice awaits further investigation.

Resistance to LPS-induced lethality has now been demonstrated in several gene-deleted mice, including macrophage migration inhibition factor (MIF 34), intercellular adhesion molecule 1 (ICAM-1 35), and TNF-RI 36, although CCR4-deficient mice are unique in that they can be resistant in both the high and low dose models of LPS-induced endotoxic shock. A great deal is known about the proinflammatory cytokine production after LPS treatment, yet relatively little about the signal transduction pathway by which LPS induces host cell activation. In the currently accepted model, LPS monomers are catalytically transferred by a lipid exchange molecule, LBP 37, to CD14 38. The LPS transmembrane coreceptor, Toll-like receptor 4 (Tlr4), interacts with the LPS–CD14 complex and initiates LPS signaling 39. The mechanism by which CCR4-deficient mice are resistant to LPS is unclear, but may be due to a difference in downstream signaling events. The use of gene microarrays may help to elucidate the differences between the CCR4^+/+ and CCR4^−/− mice in this respect.

In conclusion, we have shown that 24 h after high dose LPS treatment there is a significant reduction in the numbers of macrophages found in the peritoneum of CCR4^−/− mice. At the same time, we noted a decrease in macrophage-associated serum cytokines TNF-α, IL-1β, and MIP-1α. In addition, peritoneal lavage cells had decreased levels of mRNA for the chemokines MDC and MIP-2, which are thought to be produced mainly by activated macrophages. Taken together, these results are consistent with either a defect in macrophage function or the absence of a specific population, which is supported by the disappearance of the F4/80⁺ population at 24 h after LPS treatment. Identification of the precise mechanisms of LPS-induced cell stimulation is important for our understanding of bacterial pathogenesis and for the development of strategies to protect against gram-negative bacterial infection. Targeted deletion of the CCR4 gene has revealed an unexpected role for this receptor in LPS-induced endotoxic shock. Thus, although there is only limited relevance of the endotoxic shock models used here in mice, to the condition of septic shock in humans, it remains to be seen whether neutralization of CCR4 either by antibodies or chemokine receptor antagonists will have any therapeutic effect in vivo. Further studies with this animal model should also help elucidate the role of CCR4 in immunity, inflammation, and other biological functions.

The authors wish to thank Dr. Marie Kosco-Vilbois for critical reading of the manuscript, and Prof. Thomas Martin, Dr. Jerome Pugin, Dr. Manfred Kopf, and Dr. Irene Garcia for helpful discussions.